Proteomics is a branch of biotechnology concerned with applying the techniques of molecular biology, biochemistry, and genetics to analyze the structure, function, and interactions of the proteins encoded by the genes of an organism. The term proteomics is somewhat analogous to the term genomics in that proteomics is the study of the proteome, the entire complement of proteins in a given biological organism at a given time, while genomics is the study of the genome, the genetic make-up of an organism. Even though the proteome of an organism is the product of that organism's genome, the proteome is larger than the genome, especially in eukaryotes, in the sense that there are more proteins than genes. This is because the genome of an organism is a rather constant entity, while the corresponding proteome differs from cell to cell and is constantly changing through its biochemical interactions with the genome and the environment. A single organism will have radically different protein expression in different parts of its body, in different stages of its life cycle and in different environmental conditions. For example, results from the Human Genome Project indicate that there are far fewer protein-coding genes in the human genome than there are proteins in the human proteome (˜22,000 genes vs. ˜400,000 proteins). The large number of proteins relative to the number of genes encoding those proteins results from mechanisms such as the alternative splicing of transcripts and the post-translational modification of proteins. There is an increasing interest in proteomics, primarily because proteins are involved in virtually every cellular function, control every regulatory mechanism and are modified in disease (as a cause or effect).
Proteomics typically involves the analysis of the proteins contained in a biological sample, such as a cell lysate. Methods of analyzing the proteins in a biological sample are often grouped into two categories: top-down methods and bottom-up methods. Top-down methods involve the characterization of the intact proteins in the sample, while bottom-up methods involve breaking the proteins in the sample into a number of component peptides, and then determining the identities of the those peptides. An advantage of top-down methods is that they can identify properties of intact proteins, such as whether the protein is phosphorylated or unphosphorylated, that cannot be detected if the proteins are broken down into peptides. A disadvantage of top-down methods is that they often unable to effectively identify individual proteins. Even though bottom-down methods are more effective in identifying proteins, they cannot be applied effectively when there are a large number of proteins in the sample. Consequently, many proteomics analyses involve a combination of top-down and bottom-up methods.
In an analysis employing both top-down and bottom down components, the role of the top-down component is typically to segregate the individual proteins in a biological sample. The traditional method of segregating a mixture of proteins is two-dimensional gel electrophoresis (2-DE). In a standard 2-DE process, a biological sample containing a mixture of proteins is placed on a separation gel, which is usually a polyacrylamide matrix. The proteins on the gel are separated according to differences in a first property, typically mass, along a first direction. The proteins in the gel are then separated according to differences in a second property, typically isoelectric point, along a second direction that is orthogonal to the first direction. The two-dimensional separation process produces one or more protein “spots” distributed across a 2-D surface of the gel. Unfortunately, 2-DE methods are only effective in isolating the most abundant proteins in a sample. This lack of sensitivity primarily results from the fact that high abundance proteins mask the presence of low abundance proteins. Even if low-abundance proteins are effectively isolated from higher abundance proteins during 2-DE, the subsequent analysis of low-abundance proteins by mass spectrometry (MS) is hindered by the fact that the amount of protein required for identification by MS is near the detection limits of the most sensitive methods used to visualize the protein spots on 2-DE gels, and the fact that those gels can produce a significant amount of background during MS analysis.
Multi-dimensional column separations offer many advantages over traditional 2-DE methods, including a higher separating power, and reduced sample contamination and loss. A multi-dimensional column separation process involves sequentially subjecting a mixture of proteins to two or more separation processes to segregate the individual proteins. Those separation processes can include size exclusion chromatography, reversed phase High Performance Liquid Chromatography (RP-HPLC), ion-exchange chromatography, and capillary electrophoresis. For example, in the Ultimate™ 3000 integrated proteomics system manufactured and sold by the Dionex Corporation of Sunnyvale, Calif. (www.dionex.com), the proteins in a cell lysate are first separated by ion-exchange chromatography, producing a series of fractions, each of which containing one or more proteins. Each fraction is then subject to a RP-HPLC process, further refining the fractions. The number of proteins in each further refined fraction is typically low enough so that each of those fractions can be subject to a bottom-up analysis.
A bottom-up analysis of a protein sample requires that the proteins in the sample be broken down into a number of component peptides. The process of breaking up proteins into peptide components involves several steps, which typically include the reduction, alkylation, and digestion of the proteins. The reduction step reduces disulfide bonds within the protein. The alkylation step alkylates cysteine residues, and prevents the protein from assuming its native conformation. The reduction and alkylation steps allow the subsequent digestion step to be better controlled and more efficient. In the digestion step, proteins are broken up into peptide fragments by means of an endopeptidase enzyme that specifically cleaves the bonds between certain peptides in a protein. The most commonly used enzyme used for digestion is trypsin, which very specifically cleaves peptide bonds on the carboxyl side of lysine and arginine residues. As an example, bovine fetuin is a protein that is 341 amino acid residues in length. When bovine fetuin is digested with trypsin, it is broken up into twenty-five separate peptides that range in length from two to eighteen amino acid residues.
After a protein sample is digested, the resulting peptide fragments are subjected to analytical techniques that can enable the identification of the protein or proteins within the sample. One such process is peptide mass fingerprinting (PMF). The basic idea behind PMF is that the collection of peptides produced by digesting a protein constitutes a unique identifier or “fingerprint” of the protein. A protein's fingerprint is produced by accurately measuring the masses of the peptide fragments produced by digestion with a mass spectrometer such as MALDI-TOF or ESI-TOF. The PMF technique is described in detail in U.S. Pat. No. 5,538,897. An alternative method of analyzing a protein that has been digested is tandem mass spectrometry. A tandem mass spectrometer is capable of multiple rounds of mass spectrometry. For example, one mass analyzer can isolate one peptide fragment from many entering the mass spectrometer. A second mass analyzer then stabilizes the peptide ions while they collide with a gas, causing them to fragment by collision-induced dissociation (CID). This fragmentation process primarily gives rise to cleavage products that break along peptide bonds. Because of this simplicity in fragmentation, it is possible to use the observed masses of the cleavage products to match with a database of predicted masses for one of many given peptides. A third mass analyzer then catalogs the fragments produced from the peptides. Tandem MS can also be done in a single mass analyzer over time as in a quadrupole ion trap. A third alternative method of identifying proteins uses a combination of high-pressure liquid chromatography (HPLC) and MS. In this third method, a peptide mixture resulting from digestion of a protein mixture is fractionated by one or two steps of liquid chromatography. The eluent from the chromatography stage can be either directly introduced to the mass spectrometer through electrospray ionization, or laid down on a series of small spots for later mass analysis using MALDI.
In the current state of the art, there are a number of difficulties associated with proteomics analyses employing digestion. One difficulty is that when trypsin digestion is carried out in solution the digestion process can be extremely time consuming. For example, digestion processes carried out in a solution contained within a well of a multiwell plate often take several hours to complete. The rate of digestion can be increased by increasing the in-solution concentration of the trypsin enzyme, but high concentrations of trypsin can lead to autolysis of the enzyme, which produces large amount of contaminant peptides. Autolysis can be avoided by immobilizing the trypsin enzyme on a solid support, allowing the trypsin to be present at high enough levels so that the digestion process takes minutes instead of hours. For example, Applied Biosystems of Foster City manufactures and sells Poroszyme® cartridges, which contain trypsin immobilized on polystyrene beads. Trypsin digestion can be carried out in approximately five minutes by flowing a protein sample through a Poroszyme cartridge. A similar product is the StyrosZyme™ TPCK-Trypsin column manufactured by OraChrom, Inc. of Woburn, Mass. Although the use of cartridges containing immobilized trypsin can reduce the time it takes to digest a protein sample, it is impossible to process multiple samples in parallel using existing cartridge technology. Furthermore, existing cartridge products are not capable of performing the preliminary reduction and alkylation steps. These shortcomings of existing cartridge products hinder their integration into automated proteomics systems designed to process a large numbers of protein samples. Indeed, many currently available automated proteomics systems such as the Ultimate™ 3000 integrated proteomics system still perform reduction, alkylation, and digestion in multiwell plates, in spite of the time-consuming in-solution digestion process, because the use of multiwell plates allows a large number of samples to processed in parallel.
As an alternative to cartridges, immobilized trypsin can be placed within the channels of a microfluidic device. Microfluidic devices contain channels having at least one linear interior dimension, such as depth or radius, of less than 1 mm. It is possible to create microscopic equivalents of bench-top laboratory equipment such as beakers, pipettes, incubators, electrophoresis chambers, and analytical instruments within the channels of a microfluidic device. Since it is also possible to combine the functions of several pieces of equipment on a single microfluidic device, a single microfluidic device perform a complete analysis that would ordinarily require the use of several pieces of laboratory equipment. A microfluidic device designed to carry out a complete chemical or biochemical analyses is commonly referred to as a micro-Total Analysis System (β-TAS) or a “lab-on-a chip”. Thus it should be possible to perform several of the processes required for the analysis of a protein within a single microfluidic device. For example, researchers at the University of Alberta have designed a microfluidic device in which a protein sample is digested by flowing the sample through a packed bed of beads coated with immobilized trypsin, the component peptides produced by digestion are separated by capillary electrophoresis, and finally the separated peptides are introduced into a mass spectrometer. Wang et al., RAPID COMMUN. MASS. SPECTROM. 14, 1377-1383 (2000). Presumably the samples introduced into the microfluidic device designed by Wang et al. must have been already reduced and alkylated since the device is not configured to perform reduction and alkylation. The exclusion of the reduction and alkylation steps from the device described by Wang et al. may result from the difficulties associated with performing the three steps of reduction, alkylation, and digestion on a single microfluidic device. Among those difficulties are that those three steps take different amounts of time, require different chemical environments, and are optimally performed at different temperatures. The fact that the steps take different amounts of time makes it difficult to perform those three steps sequentially in a continuous flow manner on a single microfluidic device. Some researchers have sidestepped this issue by immobilizing a protein sample at a particular location within a channel in a microfluidic device by either binding the proteins in the sample to a derivatized surface or by using the combination of an electric field and hydrodynamic flows to hold the proteins in place. See e.g. Astorga-Wells et al., ANAL. CHEM. 76, 2425-2429 (2004). Once a protein sample is immobilized in a channel, the various reagents required for reduction, alkylation, and digestion are sequentially flowed through the channel. After those three processes are complete, the resulting peptides are allowed to exit the channel. Although immobilizing a protein sample does allow the reduction, alkylation, and digestion processes to be carried out on a single microfluidic device, the structures and equipment required to immobilize the sample increase the cost and complexity of the microfluidic device.
It is thus an object of the present invention to carry out the reduction, alkylation, and digestion processes in a microfluidic device in a manner that overcomes the previously described shortcomings. These and further objects will be more readily appreciated when considering the following disclosure and appended claims.